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Consensus guidelines for the use and interpretation of angiogenesis assays
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DOI:10.1007/s10456-018-9613-x
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Consensus guidelines for the use and interpretation of assays for the assessment and analysis of angiogenesis
Table of content
1. Endothelial cell migration assays Hellmut Augustin 2. Endothelial cell proliferation assays Judy van Beijnum
3. Tube formation Chris Hughes
4. Aortic ring assay Roberto Nicosia
5. Tip cells Reinier
6. Microvessel density + histology Peter Vermeulen
7. Intussusception Valentin Djonov
8. Lymph angiogenesis Kari Alitalo
9. Pericytes George Davis
10. Endothelial – pericyte metabolism Peter Carmeliet
11. Endothelial cell precursors Mervin Yoder
12. Microfluidics Jonathan W Song, Lance Munn
13. Zebra fish Brant Weinstein
14. CAM assay Robert Auerbach Patrycja Nowak-Sliwinska
15. In vivo matrigel plug assay Hynda Kleinman
16. Matrigel + embedded cells Joyce Bischoff
17. Oxygen Indued Retinopathy Lois Smith
18. Laser Induced neovascularization Lois Smith
19. Dorsal skin fold chamber Rakesh Jain, Dai Fukumura, Lance Munn
20. Hindlimb ischemia model Paul Quax
21. RT2 model Gabriele Bergers
22. Mouse models Robert Kerbel
Abstract
Angiogenesis is a complex process playing an important role in growth and development, as well as in a large array of different pathologies. Differences exist in the regulation of angiogenesis in different tissues and in different distinct diseases. For the study of dissected aspects of angiogenesis, a plethora of specific angiogenesis bioassays have been developed and are readily available. The study of a separated aspect of angiogenesis in a specific bioassay has many advantages but it should be realized that such approach can also have certain limitations. The current paper aims to present the various in vivo, ex vivo and in vitro bioassays for the assessment of angiogenesis and the most optimal way they should be executed. As such, this collaborative work is the first edition of a set of consensus guidelines on diverse angiogenesis bioassays, to serve for current and future reference.
Introduction
The process of angiogenesis – the formation of new blood vessels from pre-existing ones – is a hallmark of tissue expansion and remodeling in both physiological processes, such as wound healing, inflammation, ovulation and embryo development, and in various pathologies including cancer, atherosclerosis and arthritis [1-4]. Many of these conditions share similar characteristics, for example the occurrence of hypoxia, angiogenic growth factor production, basement membrane degradation, endothelial cell (EC) migration, proliferation and differentiation, and modulation of vascular support cells. However, dependent on the tissue or
disease under investigation, important details may differ considerably. Moreover, ECs in different vascular beds exhibit heterogeneity associated with the differentiated specialized functions of the tissue. For this reason it is often not possible to directly visualize the process and its molecular players in vivo. Therefore, different in vivo, ex vivo and in vitro bioassays have been developed to investigate the specific stages of the angiogenic process. However, making use of specific bioassays to study a part of the process, in order to extrapolate and understand the full process of angiogenesis inherently means doing concessions. It is therefore of extreme importance to understand the full potential of these bioassays. These assays have been instrumental in the study of vascular biology in growth and development [5-7] but also play a key role in the design and development of drugs that modulate for the treatment of many diseases [8-10]. Major examples of where the use of such bioassays are imperative are (i) the development of angiostatic drugs for the treatment of cancer and other angiogenic diseases [11,12], (ii) screening of natural compounds [13], (iii) the efforts to design combination therapies including angiogenesis inhibitors [14-19], (iv) the identification of the crucial role of lymphangiogenesis [20,21], (v) the interrelationship of angiogenesis and immunity [22-24], (vi) the development of imaging as diagnostic strategies [25], (vii) the study of resistance induction [26-28], (viii) development of compounds and strategies for the re-vascularization of ischemic injuries , and (ix) to improve the vascular fitness in aging vessels The current paper deals with the most important angiogenesis assays and aims at explaining advantages and limitations of these assays. The final goal of this paper not only confines the assessment of angiogenesis through such bioassays but also includes strategies to study angiogenesis in tissues, through means of assessing and quantifying microvessel density, vessel co-option, pericyte coverage and tip cell behavior.
1. Endothelial cell migration assays
EC migration is one of the hallmarks of angiogenesis and identifies an early step in the angiogenic cascade. Directed migration and invasion are key regulatory processes in sprouting angiogenesis. This process is characterized by cell-autonomous motility property but in some cases it acquires the features of collective migration , in which a group of cells coordinate their movements toward a chemotactic gradient and by establishing a precise hierarchy with leader and follower cells. Therefore, dissection of the molecular
mechanisms of EC migration is key to the understanding of angiogenesis and to therapeutically manipulate this process, be it with the objective to inhibit sprouting angiogenesis, e.g. in tumors, or to stimulate angiogenesis (e.g. in tissue regeneration or wound healing). A number of 2-dimensional (2D) and 3D cellular migration assays has been established as relatively simple in vitro read-outs of the migratory/angiogenic activity of EC in response to exogenous stimuli. Depending on the specific scientific question, different assays are available to quantitatively and qualitatively assess EC migration. The most widely employed cellular assays include different variations of the wound closure and the Boyden chamber assays.
1.1. Types of assays.
Cell culture wound closure assay. Lateral migration assays are performed to investigate the pro- or anti-migratory effect of compounds. Applying such assays, it is possible to determine chemokinesis (undirected migration) in response to certain compounds within the cell culture medium. However, it is not possible to determine the directed migration rate towards or away from a compound. This process is called chemotaxis and can be determined in assays supplying a gradient of the migration-inducing compound.
The cell culture wound closure assay is one of the simplest read-outs for the migratory activity of cells. It is a measure for the lateral 2D migration of EC in cell culture to test pro-migratory or anti-migratory compounds. Depending on the migratory effect of the tested substances, the assay is performed over 2 to 4 days. EC are grown to confluency in a cell culture dish and then scraped with a razor blade/pipette tip [29], allowing the EC at the wound edge to migrate into the scraped area. To really examine the motility contribution to the healing
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Commented [U1]: Williams PA, Silva EA. The Role of
Synthetic Extracellular Matrices in Endothelial Progenitor Cell Homing for Treatment of Vascular Disease. Ann Biomed Eng. 2015 Oct;43(10):2301-13. doi: 10.1007/s10439-015-1400-x. Raval Z, Losordo DW. Cell therapy of peripheral arterial disease: from experimental findings to clinical trials. Circ Res. 2013 Apr 26;112(9):1288-302. doi: 10.1161/CIRCRESAHA.113.300565.
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aging: Molecular mechanisms and potential treatments for vascular rejuvenation. Ageing Res Rev. 2017 Aug;37:94-116. doi: 10.1016/j.arr.2017.05.006. Epub 2017 Jun 1. Review. PubMed PMID: 28579130. Donato AJ, Morgan RG, Walker AE, Lesniewski LA. Cellular and molecular biology of aging endothelial cells. J Mol Cell Cardiol. 2015 Dec;89(Pt B):122-35. doi: 10.1016/j.yjmcc.2015.01.021. Deleted: . Deleted: to Deleted: the
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of endothelial sheet migration. Genes Dev. 2008 Dec 1;22(23):3268-81. doi: 10.1101/gad.1725808. Costa G, Harrington KI, Lovegrove HE, Page DJ, Chakravartula S, Bentley K,Herbert SP. Asymmetric division coordinates collective cell migration in angiogenesis. Nat Cell Biol. 2016 Dec;18(12):1292-1301. doi: 10.1038/ncb3443. Hamm MJ, Kirchmaier BC, Herzog W. Sema3d controls collective endothelial cell migration by distinct mechanisms via Nrp1 and PlxnD1. J Cell Biol. 2016 Nov 7;215(3):415-430. Haeger A, Wolf K, Zegers MM, Friedl P. Collective cell migration: guidanceprinciples and hierarchies. Trends Cell Biol. 2015 Sep;25(9):556-66. Deleted: are Deleted: aim
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mention the video-lapse techniques to analyze and quantify random motility
and to exclude the component related to cell proliferation, ECs are incubated with the anti-mitotic agent mitomycin . One downside of this method is that by scratching the cell layer the width of the scratch is difficult to control and therefore cannot be easily standardized. Moreover, wounding of the monolayer with a sharp object may scratch the surface of the cell culture dish and additionally damage the EC at the migration front. To obtain more reliable and reproducible results, fencing techniques have been developed that allow the controlled release of a confluent monolayer into lateral migration without wounding of the cells or the
underlying matrix [30]. Cells are grown as a monolayer in a culture dish containing a silicon template of defined size prior to seeding the cells. The silicon template is removed once the cells reach confluency allowing them to migrate laterally into the area previously occupied by the silicon template. This allows a simple and precise microscopic analysis of lateral migration over 2 to 4 days.
Wound healing assay connected with video-lapse microscopy allows studying in 2D dimension the role of collective migration in angiogenesis and vascular development. The use of aortic rings (see below) and that of specific microfluidic devices represent a further tool to describe this process in a 3D architecture. For instance wound healing assay exploited by single cell analysis and by using chimeric EC sheets obtained by infecting cells with different fluorescent proteins was instrumental to describe the following steps of EC collective migration: (i) in resting state ECs undergo random cell motility in the monolayer with a regulated dynamics of homotypic cell junctions; ii) the presence of cell-free space (i.e. the wound) and a chemotactic gradient result in the appearance at the sheet margin of leader cells, which are characterized by an aggressive phenotype with prominent stress fibers, ruffling lamellipodia and enlarged focal adhesions, formation of peripheral actin cables and discontinuous adherens junctions, which indicate mechanical coupling between leader and follower cells in the migrating cluster ; iii) as leaders start to migrate in the free space, a follower phenotype appears within cells of the monolayer.
Transwell cell migration assay – Boyden chamber assay. The Boyden chamber assay is a useful tool to study directed cell migration (chemotaxis) and cell invasion. It was originally introduced by Steven Boyden in the 1960’s for the analysis of leukocyte chemotaxis [31]. Today, a large number of different Boyden chamber devices (depending on individual needs) are commercially available. In this assay, one can distinguish positive chemotaxis (migration towards the attractant) and negative chemotaxis (migration away from a repellent). The assay is based on a chamber of two medium-filled compartments separated by a microporous membrane of defined pore size. In order to study cell migration, EC are placed in the upper compartment and are allowed to migrate through the pores of the membrane into the lower compartment. The chemotactic agent of interest or cells secreting chemotactic agents are present in the lower compartment. The membrane between the fluid-filled compartments is harvested, fixed and stained after a defined incubation time and the number of migrated cells on the bottom side of the membrane is determined by staining and subsequent microscopic analysis. A chemical gradient cannot be maintained for a long time. Boyden chamber assays are therefore usually limited to 2 to 6 hours. Boyden chamber assays are used to measure different types of chemotaxis, including haptotaxis, transmigration and cell invasion. Angiogenesis and transendothelial migration are special forms of haptotaxis, since the trigger for migration is not a chemokine but the presence of cell surface molecules. In this case, the insert of the Boyden chamber is coated with extracellular matrix proteins (collagens, fibronectin) on the bottom. The cells migrate along cellular adhesion sites. Transmigration describes the migration of cells, such as leukocytes or tumor cells through the vascular endothelium towards a chemoattractant. Therefore, this assay is not a measure of EC migration but for the transmigration of cells through the EC layer. Angiogenesis requires the invasion of EC through the basement membrane to form sprouting capillaries. The invasion process involves the secretion of matrix metalloproteases to degrade the basement membrane, the activation of EC and the migration across the basement membrane. This process can also be modeled in a Boyden chamber assay.
The bioactive molecules in Boyden chamber assays can be provided as recombinant molecules or by cells in the bottom chamber secreting specific factors. Manipulation of test cells (gain-of-function, loss-of-function) can
Commented [U5]: 1.
Schreier T, Degen E, Baschong W (1993) Fibroblast migration and proliferation during in vitro wound healing. A quantitative comparison between various growth factors and a low molecular weight blood dialysate used in the clinic to normalize impaired wound healing. Res Exp Med 193: 195–205.
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of endothelial sheet migration. Genes Dev. 2008 Dec 1;22(23):3268-81. doi: 10.1101/gad.1725808. Costa G, Harrington KI, Lovegrove HE, Page DJ, Chakravartula S, Bentley K,Herbert SP. Asymmetric division coordinates collective cell migration in angiogenesis. Nat Cell Biol. 2016 Dec;18(12):1292-1301. doi: 10.1038/ncb3443. Hamm MJ, Kirchmaier BC, Herzog W. Sema3d controls collective endothelial cell migration by distinct mechanisms via Nrp1 and PlxnD1. J Cell Biol. 2016 Nov 7;215(3):415-430.
Commented [U7]: : Nguyen DT, Gao L, Wong A, Chen
CS. Cdc42 regulates branching in angiogenic sprouting in vitro. Microcirculation. 2017 Jul;24(5). doi: 10.1111/micc.12372
Commented [U8]: Vitorino P, Meyer T. Modular control
of endothelial sheet migration. Genes Dev. 2008 Dec 1;22(23):3268-81. doi: 10.1101/gad.1725808
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LM, Yang HW, Tsai FC, Bisaria A, Betzig E, Meyer T. Engulfed cadherin fingers are polarized junctional structures between
collectively migrating endothelial cells. Nat Cell Biol. 2016 Dec;18(12):1311-1323.
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be introduced into the assay. Migration assays are often performed in combination with tumor cells, pericytes or fibroblasts
The real-time random migration assay
The use of video-lapse microscopy allows measuring migration not only as an end-point result, but gives information on single cell parameters , on morphological changes and on the influence exerted by specific substratum . Subconfluent ECs are plated on plastic surface coated with specific extracellular matrix proteins (i.e. collagens, fibronectin, vitronectin) allowed to adhere, and then observed with an inverted
microscope equipped with thermostatic and CO2 controlled chamber (e.g. Leica, DMi8 platform; Nikon, TE micorsope) Images of motile ECs are captured with a 5 min time interval over 4 h. Images were then processed with DIAS software (Solltech). A recent review on tracking algorithms offers a wide and comprehensive selection of the available tools to analyze cell motility. Generally data are displayed as a centroid plot showing the location of the geometrical centre of the cell as a function of time. Directional persistence was calculated by determining the ratio between the net path length and the total path length. Furthermore other parameters such as the total and net distance, the speed, the feature of turning angle can be calculated. Single cell trajectories were plotted using Matlab software and displayed in windrose graphs.
1.2 Limitations and challenges.
Standardization of techniques is one of the most critical issues to ensure the reproducibility of experimental results and one has to be aware that cellular in vitro systems represent only a surrogate of the in vivo situation. However, compared to in vivo experiments, in vitro assays are relatively simple to perform and they offer the possibility to pursue large scale screens of compounds affecting EC migration, e.g. supernatants of tumor cells. However, one has to consider the limitations of cellular assays. All the cell culture conditions have to be taken into consideration, as well as possible pitfalls of the assay itself. Pure populations of EC are required for migration assays. Human umbilical vein EC (HUVEC) are widely used for this analysis. However, they derive from a large vessel, whereas angiogenesis is driven by microvessels. HUVEC are primary cells and are only viable for a limited time. Moreover, one has to consider that cells in a culture dish change their expression profile and therefore their phenotype and behavior over time and passage. Moreover, reproducibility of scratch assays relies strongly on the initial degree of confluency [32]. Another important consideration is that EC in vivo are exposed to flow and sheer stress, which is missing in static cell culture models. The scratch assay is a straightforward cell culture assay to analyze EC chemokinesis. It does not have a high degree of sensitivity, but it is a useful tool to perform large scale screening experiments. As mentioned above, a potential drawback is the difficulty to standardize the wound areas. This could be solved as described above using silicon templates [30]. Likewise, several commercial suppliers have developed robust assays that also circumvent this problem. For example, IncuCyte ZOOM™ assays use a mechanical tool, called a WoundMaker™, to create 96 identically-sized scratches in each well. The WoundMaker™ is a 96-pin mechanical device designed to create homogeneous, 700-800 µm wide scratch wounds in cell monolayers on 96-well microplates. The device creates wounds without damaging the cells or the underlying plastic or biomatrix. Every scratch has the same
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Belyaev I, Al-Zaben N, Figge MT. Untangling cell tracks: Quantifying cell migration by time lapse image data analysis. Cytometry
A. 2017 Oct 4. doi: 10.1002/cyto.a.23249.
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Quantitative analysis of random migration of cells using time-lapse video microscopy. J Vis Exp. 2012 May 13;(63):e3585. 1: Serini G, Valdembri D, Zanivan S, Morterra G, Burkhardt C, Caccavari F, Zammataro L, Primo L, Tamagnone L, Logan M, Tessier-Lavigne M, Taniguchi M, Püschel AW, Bussolino F. Class 3 semaphorins control vascular morphogenesis by inhibiting integrin function. Nature. 2003 Jul 24;424(6947):391-7.
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dimension and is created with the same amount of pressure
(www.essenbioscience.com/en/products/incucyte). The cells are traced fully automated. Another example of such device is provided by Peira Scientific Instruments (Beerse, Belgium) together with analysis software and automated hardware [33].
The Boyden chamber assay is somewhat more delicate and requires more experience in handling. The most critical issue is the possible trapping of air bubbles in the lower and upper chambers during assembly. Air bubbles, which are trapped beneath a cell, will cause empty spaces on the filter since the cell is hindered to migrate by the bubble. For manual analysis, this may not of major importance, but it becomes relevant if an automated analysis is performed since a trapped cell is not distinguishable from a non-migrated one. It is definitely worth to invest some substantial time into the setup and troubleshooting of the assay in order to yield robust and reliable results. Therefore, it is recommended to include a checkerboard analysis in order to solidly distinguish between chemotaxis and chemokinesis effects. To this end, different dilutions of the substance are titrated in the upper and lower chamber. Equal concentrations in the upper and lower chamber should lead to the same migration behavior as in the control for a compound that strictly responds to a gradient, i.e. chemotaxis).
1.3. Concluding remarks. In summary, the lateral scratch wound assays [34] and the Boyden chamber assay are both robust and reliable systems to study EC migration. They are suitable for scale-up purposes in order to perform manual or automated large-scale compound screens. Multiple vendors provide multiple scratch and transwell assay systems. These systems offer good reproducibility and adequate throughput capacity. 2. Endothelial cell proliferation assays
Since the proliferation of ECs is a hallmark of angiogenesis, the measurement of this parameter has been broadly applied. Many regulators of angiogenesis have been identified, validated and developed based on screening for an effect on endothelial cell proliferation. ECs are among the most quiescent cells in the body, with proliferation rates approaching 0 under steady-state conditions. Only after activation, usually as a consequence of injury, inflammation or pathological processes such as malignant growth, they start to proliferate [35,36]. The ideal assay to measure EC proliferation would be rapid, reproducible, reliable and translatable and where possible should exclude inter-operator variability, e.g. through quantitative
computational readout rather than qualitative researcher-dependent observations [36]. This section presents different methods and will elaborate on problems and pitfalls.
2.1. Types of proliferation assays. A number of different approaches to address cell proliferation have been developed in the last decades. In general, these include the monitoring of cell number, the detection of DNA synthesis by incorporation of labeled nucleotide analogs, measurement of DNA content, detection of proliferation markers and metabolic assays (Fig. 1). Depending on the broadness of the definition of cell proliferation, which can range from the narrow description “the fraction of cells dividing over time” to the more general “the doubling time of a population over time”, different assays may be pursued. Next to that, means and equipment available will also dictate the choice for a particular method. As all methods focus on a particular aspect of the process, it is highly recommended to verify results with a complementary assay. Cell counting. Cell counting can be considered the gold standard for proliferation. Moreover, at least in theory, it is one of the most straightforward procedures of measuring proliferation of a cell population. It can be done using automated cell counters (e.g. Coulter) or by using a hemocytometer after removal of the cells from the culture vessel [36,37].
More recently, different automated platforms have entered the market that allow analysis of cells while present in microplates, such as plate cytometers, automated microscope or high-content screening platforms, that are compatible with cell counting-like procedures. With these, cells can be monitored over time but frequently require staining for detection and (computation-assisted) quantification, by e.g. staining of nuclei. More recently, real-time cell analysis (referred to as RTCA) platforms have emerged, that allow label-free, automated, real-time monitoring of cellular properties during incubation based on electrical resistance measurements. As such equipment requires considerable investment, it will be beyond reach for many laboratories.
DNA labeling. During S-phase of the cell cycle, DNA is synthesized and subsequently divided between the daughter cells (2N→4N→2N). Addition of modified nucleotides to the culture medium will result in incorporation of these into the newly synthesized DNA. Adhering to the narrow definition of proliferation as stated above, this type of assay most closely reflects a means of measuring the fraction of actively dividing cells. It should be realized that this technique does not directly measure cell division or population doublings, but exclusively only incorporation into DNA.
The use of 3H-thymidine, has a long track record [35,37,38,18]. Briefly, cells are pulsed with 3H-thymidine for
several hours and radioactivity is measured by liquid scintillation counting. It provides a very accurate representation of actively proliferating cells and is highly sensitive since the amount of incorporated 3
H-thymidine is directly proportional to the rate of DNA synthesis [36,37]. Constraints on using radioactive compounds and the rise of alternative methods have limited its use somewhat nowadays. In a similar approach, the incorporation of 5-bromo-2’-deoxyuridine (BrdU) or EdU (5-ethynyl-2’deoxyuridine) can be measured. BrdU or EdU can be (in)directly detected and subsequently be (semi-)quantified using ELISA, flow cytometry or immunohistochemistry [36,37,39], the latter two quantification techniques allowing to determine the fraction of dividing cells. These uridine analogs can be combined with DNA dyes (see below) to gain additional cell cycle information [2].
A more static approach is the measurement of cellular DNA content using intercalating dyes such as PI (propidium iodide) or DAPI (4',6-diamidino-2-phenylindole). Using flow- or plate cytometry, a profile of the distribution of cells over the different phases of the cell cycle can be visualized, represented by DNA contents of 1N (G1/0), 2N (G2/M), or mixed (S). In addition, this method allows for the detection of apoptotic cells that would exhibit a subG1/0 (<1N) DNA content.
An alternative method to study EC cycle is based on the use Fucci (fluorescent, ubiquitination-based cell cyle indicator) technology (Termofisher). It consists of a fluorescent protein-based system that employs both a red and a green fluorescent protein respectively fused to cdt1 and geminin, which are two regulators of cell cycle. These two proteins are ubiquitinated by specific ubiquitin E3 ligases in a specific temporal sequence. In the G1 phase, geminin is degraded; therefore, only cdt1 is present and appears as red fluorescence within the nuclei. In the S, G2, and M phases, cdt1 is degraded and only geminin remains, resulting in cells with green fluorescent nuclei. During the G1/S transition, when cdt1 levels are decreasing and geminin levels increasing, both proteins are present, giving a nuclear yellow fluorescence. More recently, Fucci probe was re-engineered to generate a triple color-distinct separation of G1,S and G2 phases extending the use of this technology to quantitative analyze the interphase of cell cycle
Proliferation markers. Cell division is a highly coordinated process where specific proteins show a concerted action to allow mitosis to complete. Detection of these proteins, usually through immunochemical procedures, allows the estimation of the fraction of dividing cells. This approach can be used in in vitro end-point assays, but can additionally be used to evaluate active EC proliferation in tissue sections (see separate section).
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N, Hiratsuka T, Kogure T, Hoshida T, Goshima N, Matsuda M, Miyoshi H, Miyawaki A. Genetically Encoded Tools for Optical Dissection of the Mammalian Cell Cycle. Mol Cell. 2017 Oct 25. pii: S1097-2765(17)30750-5
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Metabolic assays. Gradually, the use of cell viability assays has taken a rather dominant position in addressing cell proliferation. While not reflecting this property in its narrowest sense, if properly conducted they accurately represent the number of live cells in an assay system. They are readily available and require minimal handling and infrastructure. The most well-known is the MTT assay (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide), in which this yellow salt is taken up by metabolically active cells and converted by mitochondrial dehydrogenase to insoluble purple formazan crystals that cannot leave the cell. As the amount of the converting enzyme is highly stable in a given cell population, the formation of formazan (and hence color intensity) is proportional to the number of viable cells. This is subsequently quantified by solubilization of the crystal containing cells and spectrophotometry. Variations to this method, e.g. involving less toxic reagents, simplified reaction steps or alternative readouts such as cellular ATP levels, have also been widely used [37,33,40].
2.2. Limitations and challenges.
Endothelial cell culture considerations. Studying EC proliferation in vitro requires a purified population of ECs compatible with the assay setup. HUVEC are a widely available source, but are of macrovascular origin, whereas angiogenesis in vivo takes place at the microvascular level. Therefore, other sources of ECs are necessary for confirmation of results. Foreskin-derived human dermal microvascular EC (HDMEC) are therefore a good alternative.
Like with all primary isolates, cells have a limited life span in vitro. Moreover, their mere propagation in culture induces phenotypic changes [36,37]. As such, it is recommended to only use populations of cells that underwent limited population doublings. Though immortalized EC can pose a helpful alternative, it should be recognized that the immortalization itself may or will likely alter growth control and survival mechanisms in these cells [41]. As such, care must be taken to address the generalizability of assay outcome.
In all cases, cell density needs to be carefully controlled. Assay linearity can be compromised when cells are plated too dense (e.g. 50,000 cells/cm2) or too sparse (e.g. 5,000 cells/cm2). Loss of cell-cell contact is a
potent stimulus for EC to proliferate, whereas EC enter a quiescent state upon confluency, known as contact-inhibition [36,37,42]. Optimization of the dynamic performance can involve synchronization of cells by exposure to low serum conditions (when studying pro-angiogenic molecules), or by stimulation prior to the addition of anti-angiogenic drugs [6].
Assay choice considerations. Each type of EC proliferation assay described here has its own limitations. Though cell counting is the most straightforward method, it can be prone to sampling error when cell detachment is required. Furthermore, it can be labor intensive and requires relatively large samples [36,37]. However, it generally does not require the handling of toxic, mutagenic or radioactive compounds like with several metabolic, DNA labeling and DNA incorporation based assays [36,37,42].
From a methodological point of view, each assay has its strengths and weaknesses. For example, the indirect detection of antigens (e.g. PCNA or BrdU) requires careful procedural optimization. For the latter, the alternative ‘click’ chemistry by which the analogous EdU can be detected directly, this issue is circumvented [37,36,39]. In addition, assay readout and interpretation are important to consider. When measuring incorporation of nucleotide analogs, one should realize that DNA synthesis is not in all situations confined to chromosomal duplication during S-phase [37,36]. For example, during DNA repair nucleotides are excised and replaced, which is especially relevant when addressing the action of compounds with a potential DNA damaging effect. With the DNA intercalating dyes, care must be taken that duplets are excluded in the gating procedure or with readout in plate-based systems. By nature, this type of assay is mostly suited for truly diploid cells, and not for cells that may display alternative karyotypes. Although the latter is not a common trait of EC, a few reports have addressed this matter in tumor-derived EC [43], and personal observations also indicate
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this may be the case with EC lines. Finally, test reagents may interfere with readout chemistry, e.g. compounds that affect mitochondrial function are less compatible with metabolic assays.
2.3. Concluding remarks. The choice for a particular EC proliferation assay is determined by a number of considerations. Endpoints, test compounds, laboratory infrastructure, scale, required throughput, convenience and cost all influence the applicability of an assay system. The growth of a cell population is influenced by both division and death, which is difficult to simultaneously monitor. Most important is the researchers’ awareness that all assays have their strengths and weaknesses and that interpretation of data is done with care, and if possible, validated with an alternative method.
3. 3D models of vascular morphogenesis
To generate blood vessels and vascular networks, ECs must undergo vascular morphogenesis, which can include either vasculogenic and/or angiogenic processes [44]. In vasculogenesis, differentiating ECs migrate, proliferate, aggregate and rearrange to form cords that then undergo lumen formation to generate three-dimensional (3D), tubular blood vessels. In angiogenesis, stimulatory vascular endothelial growth factor (VEGF) promotes EC sprouting from existing vessel walls, the formation of new tubes, and anastomosis (joining) with other vessels to create an interconnected network of vessels. ECs recruit perivascular stromal cells (pericytes) to stabilize this newly-formed network and minimize leak upon blood perfusion. Importantly, not all sprouts become functional vessels. Pruning serves to selectively remove redundant or non-functional vessels to optimize fluid flow through the network [45].
In vitro assays have played a valuable role in our understanding of vascular morphogenesis. These assays provide a simpler platform than animal models for dissecting individual steps within the process while also incorporating 3D matrix to mimic native in vivo tissues. Here, we present several of the most reliable and informative assays developed to date and highlight the strengths and limitations of each (Table 1). While many types of ECs can be used in these assays, the most commonly used are HUVECs and human endothelial colony-forming cell-derived EC (ECFC-ECs), which generally have a higher proliferative potential. Mouse ECs are not generally used in these assays as they are notoriously hard to maintain in culture. While we use “ECs” to reference both cell sources here, assays using a specific EC source are annotated accordingly.
3.1 Types of assays.
Fibrin Bead Assay. Traditional Matrigel cord-forming or collagen I angiogenic invasion assays are insufficient to model the complexity of angiogenesis, as these assays are two-dimensional and ECs in these assays often form incomplete lumens. Moreover, lumen formation in Matrigel is not unique to ECs as several non-EC cell types (e.g. human prostate carcinoma and glioblastoma cells) also form lumens, complicating the interpretation of results from these assays [36]. In contrast, the fibrin bead assay provides a platform for testing EC sprouting and lumen formation over an extended period (2-3 weeks), incorporates a 3D, extracellular matrix (ECM), and multiple cell types (i.e. stromal pericytes) to model native angiogenesis. ECs (HUVECs) are first allowed to adhere to collagen I-coated Cytodex beads to generate an EC monolayer that mimics the vessel wall of native vessels. These EC-coated beads are then embedded into a fibrin gel with human stromal cells either embedded within the gel or plated in monolayer on top. Tip cells are observed 2-3 days post-plating and elongating sprouts appear 2-4 days after this (Fig. 2a-b). When maintained in pro-angiogenic EGM-2 medium (Lonza), lumens form within a week and the assay remains viable up to three weeks, at which point
anastomosis between sprouts is often apparent [46]. A detailed, video protocol of this assay is available [47]. Angiogenic sprouting from individual beads is evaluated by phase-contrast microscopy allowing for
quantification of sprout number, length of sprouts, percentage of sprout lumenization, and the number of anastomoses. Genetic approaches (siRNA, lentiviral transduction) [48] can modify gene expression in
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individual cell types to dissect cell-autonomous components of the angiogenesis process. Protein expression and localization is measured by fixing bead assays and using modified immunofluorescent staining techniques. More detailed gene expression analyses are made possible by harvesting individual cell types to track RNA expression changes over time through various stages of sprouting angiogenesis.
The use of stromal cells (i.e. lung fibroblasts) is critical to the success of this assay, as these cells secrete angiogenic factors necessary for EC sprouting and lumen formation, including growth factors (e.g. HGF, TGF-α, and Ang-1), as well as ECM molecules, matrix-modifying proteins and matricellular proteins (e.g. collagen I, procollagen C endopeptidase enhancer 1, secreted protein acidic and rich in cysteine [SPARC], trans-forming growth factor-β–induced protein ig-h3 [βIgH3], and insulin growth factor–binding protein 7 [IGFBP7]). These factors act to locally stiffen the matrix, which supports sprouting and lumen formation [49]. This assay represents a significant improvement over conventional, single cell-type angiogenic assays, as the inclusion of multiple cell types more closely mimics the physiological environment. Nevertheless, as this assay uses primary cultures of cells, rather than cell lines, it is important to remember that batch-to-batch variations in stromal cells (and HUVECs) can significantly affect assay results. To partially overcome this issues, it should be appropriate to use ECs pooled from 5-10 umbilical cords. Generally, for this assay to be reproducible, it is crucial to identify stromal cell-HUVEC pairs that yield optimal angiogenic sprouts for this assay to be reproducible.
Collagen Lumen Assay. To understand EC lumen formation mechanics, early assays seeded ECs in monolayer on plastic dishes coated with ECM proteins (i.e. collagen I, collagen III, fibrin, or Matrigel). While these 2D assays are sufficient to induce EC cord formation [50-52], they cannot reproduce the necessary cues for true lumen formation found in native, 3D tissues. Collagen sandwich assays surround ECs within a 3D matrix by seeding the cells in monolayer on collagen I matrix, then covering them with a second layer of collagen [53]. Nevertheless, tube formation fails to occur in a random, 3D growth pattern, forming only in the X-Y plane of the initial gel layer and not in the z-axis. As this does not adequately recapitulate normal vessel growth in a true 3D environment, George Davis and others further optimized these assays, opting instead to embed single ECs (HUVECs) randomly throughout a collagen I matrix. In the simplest version of these assays, HUVECs are seeded at low density (7x105 cells/mL) under serum-free growth conditions and with the addition
of minimal growth factors (phorbol ester, VEGF, and fibroblast growth factor-2 [FGF-2]). After 48 hours, the embedded ECs form intact tubes throughout the gel, with clearly demarcated lumens (Fig. 2c-d). Several variations on this assay have since enhanced and optimized lumen formation. First, the addition of several other growth factors, including stem cell factor (SCF), IL-3, stromal-derived factor-1α (SDF-1α), and FGF-2, further promote lumen formation while maintaining serum-free growth conditions. Second, when
simultaneously seeded within the same matrix, stromal pericytes are recruited by ECs, recapitulating a key step in vascular morphogenesis. Lastly, to understand the process of EC sprouting and angiogenesis, ECs can be seeded on top of a 3D collagen gel containing the same growth factors and invasion of the underlying gel layer can be quantified. A detailed protocol of the collagen lumen assay and its variations is available for further reading [54].
Real-time imaging of tube formation can be achieved using fluorescent protein-transduced ECs. Alternatively, fixed vessels can be stained with 0.1% toluidine blue and imaged using brightfield microscopy (Fig. 2d). More in-depth analyses can be carried out on these fixed vessels using immunofluorescence staining of relevant protein markers or transmission electron microscopy to resolve structural details of formed lumen and remodeled ECM.
Regular users of collagen gels will note that the viscosity, pH, and contraction of these gels can hinder successful execution of assays in the hands of new users. As a result, special care should be taken when pipetting (such as when mixing cells and growth factors) and plating gels to ensure even gel coating of the
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cell responses can vary from cord to cord, I suggest to use pool of HUVEC isolated from 5-10 different cords. occurring A general suggestion on the use of HUVEC and the differences within different cords is to Deleted: interleukin-3 ( Deleted: )
bottom of the well plate. Perhaps most significantly, early gel contraction can limit the useful length of these assays. Users will note that plating gels only in wells within the center of the 96 half-area wells and adding medium or water to the outer wells of the plate will minimize gel contraction, by maintaining local humidity levels. Additionally, seeding fewer ECs within the collagen I 1.5x103 cells/ml) can minimize gel contraction and
prolong the assay.
Retinal Explant Assay. Although in vitro assays are high-throughput and can mimic major steps in vascular morphogenesis, these assays do not fully recapitulate the in vivo, whole-organ environment [46]. Several in vivo animal models, such as mouse retina or zebrafish fins, are valuable tools for studying vascular (re)establishment in a physiologically-relevant context [36,55]. However, the added complexity of these systems makes it more difficult to ascertain the role of individual proteins and growth factors in the vascular morphogenesis process, relying on genetic manipulations or system-wide administration of pharmacologic inhibitors to dissect molecular pathways [36,56]. As such, there is a need to increase assay complexity (and physiological relevance) while developing platforms amenable to ex vivo study in the laboratory. Retina explant assays are one such ex vivo platform, whereby dissected retinas are maintained and observed for vascular morphogenesis over several weeks in the laboratory. While multiple versions of this assay have been published, a protocol published by Sawamiphak, et al., is most widely used for the study of endothelial sprouting [57]. Briefly, retina cups from embryonic, postnatal, or adult mice are harvested and cut radially to allow flat mounting of the retina interior surface onto a membrane insert. After recovery in media for 2-4 hours, the explants can then be treated with stimulatory or inhibitory agents for up to 4 hours, followed by whole-mount microscopy analysis to evaluate the (anti-) angiogenic effect of these agents on vessel sprouting (Fig. 2c). When perfected, each pair of retinas can be harvested and dissected within minutes. Unfortunately, without the support of a 3D matrix, retinal cells cannot survive for long periods, thus making studies for later stages of angiogenesis impossible. To overcome this, Rezzola, et al., have improved the assay by embedding the retinas in different matrices after dissection [58]. In this approach, retinas can be cross-cut into 4 equal pieces and left in serum-free media overnight. The retina fragments are then embedded in Matrigel, collagen I, or fibrin matrix and fed every 2 to 3 days. Depending on the age and the matrix used, sprouts can be observed between day 3 to 6 and anastomosis of neighboring sprouts, similar to what occurs in vivo can be observed in 10-14 days [59] (Fig. 2d). These explants can be maintained up to 3 weeks before the vessels eventually regress.
Vessel formation can be analyzed in real time using time-lapse imaging or the explants can be fixed and imaged by immunofluorescent microscopy at set experimental time points [59,60]. Gene expression can be manipulated by genetic crossing of the donor mice or, more transiently, by treating retinas with lentivirus or siRNA. Moreover, embedded retinas can be treated with drugs over extended periods to dissect individual signaling pathways.
Several factors are critical to consistently achieve sprouting from dissected retina explants. First, the matrix proteins in which retinas are embedded can greatly influence how vessels sprout. In our experience, Matrigel is far superior to single matrix proteins in inducing sprouting. However, the addition of 10-20% Matrigel in collagen I matrix is sufficient to stimulate sprouting compared to pure collagen I matrix. Second, the use of pro-angiogenic EGM-2 yields more sprouts as compared to basal medium alone. Lastly, as with any tissue explant, the age of the mouse can influence the degree of vessel sprouting. As such, special care should be taken to select mice appropriate for the experimental question at hand. There are many similarities between the mouse retinal explant assay and the traditional mouse/rat aortic ring angiogenesis assay or rat vena cava explant assay [61-63]. However, retinal explant models more closely model true capillary sprouting as the vasculature in these explants is actively developing and remodeling. This makes the retinal explant model uniquely suited to studying microvessel formation and its underlying mechanisms.
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Vascularized Micro-Organ (VMO) Platform. To understand all the steps of vascular morphogenesis in a single platform, our group has optimized a vascularized micro-organ (VMO) approach to drive formation of a perfusable vascular network within a 3D hydrogel matrix environment. In contrast to the assays described above, VMO-embedded ECs are exposed to and respond to shear stress, form lumenized vessels, and are perfused with a blood-substitute medium that delivers nutrients to tissues within the matrix, just as in the body. Specifically, this platform utilizes “arteriole” (high pressure) and “venule” (low pressure) microfluidic channels that are joined by a living microvascular network that forms by vasculogenesis in an intervening tissue chamber (Fig. 2g). A pressure differential between the two channels is used to drive interstitial flow through the fibrin gel matrix during vessel development, induce vessel formation (through shear-sensing), and drive convective flow through the mature vasculature once formed and anastomosed to the outer channels. This pressure difference is induced by varying the level of medium within fluid reservoirs at either end of the microfluidic channels, thereby creating hydrostatic pressure heads that ensure continuous fluid convection across the cell chamber. To form vessels, human ECFC-ECs and human lung stromal cells are co-loaded within a fibrin matrix into the central cell chamber through an independent loading tunnel. When maintained in pro-angiogenic EGM-2 medium, vessels form within 4-6 days post-loading (Fig. 2h). When perfused with 70kDa rhodamine-dextran, a molecule similar in size to albumin, these vessels demonstrate minimal vessel leak – comparable to in vivo microvasculature. For readers interested in more information, a detailed protocol for loading and maintaining the VMO platform is available [64].
The VMO platform is fabricated from polydimethylsiloxane (PDMS), an optically clear, biologically inert polymer widely used in the microfluidics field [65]. The use of this polymer and the dimensions of the platform ensure that live, GFP-transduced EPCs can be imaged and quantified throughout vessel formation. Specific parameters such as vessel network length, branching, and anastomosis can be measured in real-time as can vessel permeability by perfusion with fluorophore-tagged dextran molecules of various molecular weights. Additionally, immunofluorescent staining can be used to quantify expression of specific molecular markers or RNA can be collected to measure changes in gene expression. Lastly, gene expression can be manipulated by treating individual cell types with lentivirus or siRNA prior to loading in the platform.
To ensure robust and reproducible vascular network formation, several steps are critical. First, the fibrin gel matrix must be consistently loaded within the VMO cell chamber. During normal loading, perfusion burst valves at the interface between the tissue chamber and the microfluidic channels ensures a gel/air interface (later a gel/fluid interface) is formed. To simplify loading and minimize specialized training for new users of the platform, current iterations of the VMO platform incorporate a pressure release valve at the loading tunnel that minimizes unintended gel bursting [66]. Second, robust vascular network formation requires that vessels within the chamber anastomose with the outer microfluidic channels. To facilitate the formation of these
anastomoses, ECs can be either seeded directly within the microfluidic channels or induced to migrate from the gel by coating the external channels with extracellular matrix [67]. As with the fibrin bead assay, optimal stromal cell-EC pairs should be validated to ensure assay reproducibility.
3.3. Limitations and challenges. With all EC assays, the source of ECs is critical to assay success. Although commercial versions of HUVECs and ECFC-ECs are available, cells from these sources show limited utility in many 3D assays, likely due to a larger than optimal number of cell doublings prior to shipment and use in the laboratory. As a result, the use of primary-isolated ECs will provide the most consistent results and is strongly encouraged. Readers will note that isolation protocols for both cell types are available [68,69]. Additionally, patient-to-patient variation between different EC isolations can lead to inconsistent assay results, an issue that may be avoided by pooling several EC lines prior to use.
Given the many differences between the assays described here, the useful length of these assays varies considerably. Even in well-trained hands, the contraction of gels in the collagen lumen assay effectively limits the useful time frame of assays to 72 hours or less. However, the other assays described here can persist for much longer periods of time, with the fibrin bead assay, retinal explant assay, and VMO platform all suitable for time points up to three weeks under appropriate conditions.
Lastly, the majority of these assays can be run in a relatively high-throughput manner, thereby accelerating the speed with which genetic, molecular, or pharmacologic screens can be conducted to understand vascular morphogenesis. This is especially true with the fibrin bead and collagen lumen assays, which utilize multiple beads or multi-well culture plates to increase assay throughput. Similarly, while initial versions of the VMO platform were cumbersome to load in high-throughput numbers, this platform is now used in an optimized configuration that incorporates up to 16 individual VMO devices within a standard 96-well plate [70]. This design simplifies translation to outside labs and interfacing with existing microscope and plate reader
infrastructure. Of all the assays described in this section, retinal explants are most adversely affected by delays between initial dissection, mounting, and plating of tissue samples. This inherently limits the number of animals that can be dissected at once and, for now, limits the number of retinas that can be screened simultaneously. 4. Aortic Ring Assay
Explants of rat aorta have the capacity to sprout and form branching microvessels ex vivo when embedded in gels of extracellular matrix. Angiogenesis in this system is driven by endogenous growth factors released by the aorta and its outgrowth in response to the injury of the dissection procedure. This property of the aortic wall first described in the early 1980s [71], led to the development of the aortic ring assay [72] which is now widely used to study basic mechanisms of angiogenesis and test the efficacy of proangiogenic or antiangiogenic compounds [73].
4.1. Benefits and strengths of the Aortic Ring Assay. The aortic ring assay offers many advantages over existing models of angiogenesis. Unlike isolated ECs, the native endothelium of the aortic explants has not been modified by repeated passages in culture and retains its original properties. The angiogenic response can be inhibited or stimulated with angiogenic regulators and analyzed by molecular or immunochemical methods without the confounding effects of serum. Angiogenic sprouting occurs in the presence of pericytes, macrophages and fibroblasts, as seen during wound healing in vivo [73]. The different cell types can be identified with specific cell markers by immunostaining whole mount preparations [74] of the aortic cultures. The ultrastructure of neovessels at different stages of development can be evaluated by electron microscopy (Fig. 3). Many assays can be prepared from the thoracic aorta of a single animal (approx. 20-25 cultures/rat aorta; approx. 10-15 cultures/mouse aorta). The angiogenic response can be quantitated over time, generating curves of microvascular growth. Aortic cultures can be used to study mechanisms of vascular regression, which typically follows the aortic angiogenic response as seen during reactive angiogenesis in vivo. Aortic rings transduced with viral constructs or obtained from genetically modified mice can be used to study the role of specific gene products in the regulation of the angiogenic response [73].
Recently, rat aortic ring assay was adapted to human arteries by using matrigel as 3D hydrogel. Arteries are maintained in EC basal medium (EBM / MCDB131) supplemented with 5% fetal calf serum and heparin. When growth factors are added (VEGF-A, EGF, FGF-2), capillary outgrowth begins from around day 8 to 12 and continues to develop up day 30. As reported for rat aortic ring, this assay is suitable to gene editing by gain- and loss-of- function approaches and for drug screening
4.2. Assay overview. A detailed description of the aortic ring assay protocol is available in previous reports [75,76]. We provide here a summary of key steps for the preparation of the assay. Aortic rings are prepared from the thoracic aorta of 1-2 month old rats or mice. After excision from the animal, the aorta is transferred to
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PA, di Blasio L, Sessa R, Bussolino F, Primo L. Modeling human tumor angiogenesis in a three-dimensional culture system.
Blood. 2013 May 23;121(21):e129-37.
a Felsen dish containing serum-free EBM . Under a dissecting microscope, the aorta is cleaned of blood and fibroadipose tissue using Noyes scissors and microdissection forceps. Care is taken not to stretch, cut or crush the aortic wall during the isolation and dissection procedures. As the dissection progresses, the aorta is rinsed in the four compartments of the Felsen dish. Using a scalpel blade, the aortic tube is then cross-sectioned into 0.5-1 mm long rings. The proximal- and distal-most rings, which may have been damaged during the dissection procedure are discarded. The remaining rings are washed through sequential transfers into eight consecutive baths of serum-free medium, using compartmentalized Felsen dishes. Aortic rings are then embedded individually into thin collagen, fibrin or basement membrane gels as described. Once the gel has set, 500 μl of serum-free EBM is added to each culture. Each experimental group comprises quadruplicate cultures in 4-well NUNC dishes. Aortic ring cultures are incubated in a humidified CO2 incubator at 37°C for
7-21 days.
4.3. Quantitative Analysis of Angiogenesis in Aortic Cultures. The angiogenic response of the rat aorta can be quantitated by visual counts or by computer-assisted imaging. For visual counts, cultures are examined every 2-3 days and scored for angiogenic sprouting by using an inverted microscope with bright-field optics equipped with 4X to 10X objectives and a 10X eyepiece. Angiogenesis is scored by counting microvessel sprouts, branches and loops according to previously published criteria [72]. Aortic outgrowths can be also quantified by image analysis using low power images of the cultures thresholded to highlight the vascular outgrowths [77-80]. Standard statistical methods are used to analyze data and determine levels of significance between control and treated cultures. An internal control group with untreated aortic rings must be included in each experiment to mitigate the effect of possible interassay variability.
4.4. Critical points. For this assay, we recommend using the thoracic aorta because of its uniform size and intercostal artery branching pattern. The abdominal artery can also be used, but its variable pattern of collaterals and tapering lumen may introduce variability in the angiogenic response. Injury to the aortic endothelium may be an additional cause of uneven sprouting from different rings. Therefore, special care must be taken not to damage the aorta by stretching or letting it dry during the isolation and dissection procedures. Dissection of the aorta and preparation of the aortic ring cultures are best performed in a tissue culture room with HEPA-filtered air to avoid microbial contamination. Best results with this assay are obtained using interstitial collagen or fibrin gels. Collagen can be produced in-house, as described [75,76], or purchased from commercial sources [81]. Fibrinogen and thrombin for the fibrin gel are commercially available. Matrigel, a basement membrane-like matrix of tumor origin, can also be used [82]. Matrigel cultures, however, require growth factor supplements due to the limited ability of the aortic rings to sprout spontaneously in this dense matrix. The growth medium used for the assay should be optimized for the growth of ECs in the absence of serum. Optimal results in collagen and fibrin cultures can be obtained with EBM. When preparing individual collagen gel cultures, given the small volume of gel (20-30 µl), it is important to remove excess growth medium from the aortic rings when they are transferred into the collagen, fibrinogen or Matrigel solution. This is accomplished by gently streaking the aortic ring onto the bottom of the culture dish while holding it from the adventitial side with microdissection forceps. When working with fibrin gels, which set rapidly, no more than four cultures at a time should be prepared, to avoid disrupting the developing gel while positioning each ring. In addition, for fibrin cultures, the culture medium should include a plasmin inhibitor such as epsilon aminocaproic acid (EACA) to inhibit fibrinolysis by the aortic rings, which would rapidly destroy the matrix needed for ECs to sprout.
The rat aortic ring assay is robust and very reproducible when performed by an experienced operator. The mouse aortic ring assay is more variable than the rat aortic ring assay, likely because of the small size of the rings. For this reason, at least twice as many aortic rings should be used for this assay. Miniaturization of the
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Commented [U19]: I suggest to avoid this
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assay using IBIDI microchambers and a smaller volume of growth medium (50 µl) is recommended for the mouse aortic ring assay to ensure spontaneous sprouting under serum-free conditions [76]. In all cases, experiments should be repeated 2-3 times to obtain sufficient number of data points for statistical analysis. The growth medium can be replaced on a regular basis (3 times/week) or left unchanged for the duration of the experiment. If the medium is not replaced with fresh medium, the angiogenesis response and the stability of neovessels are enhanced due to accumulation of endogenous growth factors in the system. For
immunohistochemical evaluation of the aortic cultures, biomatrix gels should not exceed 20-30 µl and should be well spread as a thin wafer around each ring. Formalin fixation should be limited to 10 min to avoid excessive cross-linking of proteins. In addition, overnight incubation may be needed for optimal penetration of the primary antibody into the gel.
4.5. Limitations and challenges. The main limitation of the aortic ring assay is the lack of blood flow, particularly for angiogenesis-related genes that are regulated by mechanochemical mechanisms. An additional potential limitation is the source of angiogenic ECs, which are arterial and not venous, as neovessels in vivo primarily sprout from postcapillary venules. Many studies performed with this assay, however, have shown good correlation of results obtained with the aortic ring assay and in vivo models of angiogenesis. If needed, the aortic ring assay methodology can be applied to veins as reported [63]. Some investigators have described variability of the angiogenic response in different aortic cultures. This is due to the delicate nature of the endothelium which can be damaged because of inadequate handling of the aorta or the aortic rings, drying of the explants, or excessive exposure of these to alkaline pH. Suboptimal preparation of the gels resulting in a defective matrix scaffold can also result in a poor angiogenic response. In addition, the age and genetic background of the animal significantly affect the capacity of the aortic rings to sprout spontaneously or in response to angiogenic factors. Aortic outgrowths in Matrigel are much denser than in collagen and fibrin and more difficult to quantitate by visual counts due to the intricate branching pattern of the endothelial sprouts and the tendency of mesenchymal cells to arrange in confounding networks, which mimic angiogenic sprouts. Immunostaining of the aortic outgrowths with endothelial markers followed by image analysis may overcome this limitation. For quantitative analysis of the angiogenic response, the visual count method (described in detail in reference [76]), becomes challenging when cultures stimulated by growth factors produce 250-300 or more vessels. Since the outgrowths of rings oriented with the luminal axis parallel to the bottom of the culture dish (recommended orientation) are typically symmetrical, angiogenesis in these cases can be quantitated by counting the number of microvessels in half of the cultures and then doubling the score. Alternatively, these cultures can be measured by image analysis [77-80]. Finally, for the whole mount immunohistochemical stain, gel thinness is critical for optimal antibody penetration.
4.6. Concluding Remarks. Many of the molecular mechanisms orchestrating angiogenesis have been discovered, but many others remain to be identified, studied and evaluated as targets for the development of new therapies. The aortic ring assay reproduces ex vivo cellular and molecular mechanisms that are essential for the regulation of the angiogenic process. As such, this assay provides invaluable testing ground to test new hypotheses and analyze the efficacy of the next generation of angiogenesis-targeting drugs.
5. Detection methods for tip cells
The tip cell is the first specific angiogenic EC phenotype that becomes activated during angiogenesis, and forms the leading cell on the tip of a vascular sprout. Tip cells are characterized by their position, their long and dynamic filopodia, absence of proliferative activity and their migratory behavior. Stalk cells that follow the tip cells have other properties, they proliferate, produce ECM, and form a vessel lumen. Recently, the tip cell has been studied to decipher the functional relevance of its specific morphology, its coordinated behavior and its
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distinct gene expression profile. Studies in mouse- and zebrafish development, in in vitro sprouting assays and in tumor angiogenesis have illustrated that VEGF and Notch signaling are essential for the differentiation of tip and stalk cells. However, a more detailed understanding of the underlying mechanisms of tip cell differentiation is still lacking, especially due to its dynamic phenotype, and the absence of specific and definite in vivo markers for tip cells.
5.1. Tip cell detection methods. Tip cells have mostly been studied in vivo, i.e. in the developing mouse retina and in zebrafish, and to a lesser extent in vitro in angiogenic sprouting models. The basis for tip cell research was set by Gerhardt et al. 2003 [83], who described the tip cell as a specialized EC that could be distinguished from other EC phenotypes (stalk cells, phalanx cells and quiescent ECs) and who provided a comprehensive overview on how to detect these cells by tip cell specific markers using immunostaining or in situ hybridization. In the search for additional tip cell-specific markers whole genome genetic profiling strategies were used [84,85]. In recent years, more and more transgenic models are increasingly being used. Fig. 4.
Labeling of tip cells in vivo. The most extensively used model for tip cell research is the developing mouse retina. Here, the first vessels originate from the optic nerve and grow radially to the peripheral retinal margin during the first week after birth. Subsequently, the superficial capillaries start sprouting downwards to form the deep plexus and the intermediate vascular plexus, and in approximately the third postnatal week all vascular layers are completely mature. Sprouting angiogenesis can ideally be studied in the first postnatal week, when tip cells are located in the angiogenic front.
Immunofluorescent staining of retinal vasculature with biotinylated isolectin B4 (IB4) labels sugar (α-d-galactosyl) residues on the endothelium and endothelial tip cell filopodia, but also marks microglia and macrophages. Other useful antibodies against surface bound proteins are VEGFR2, VEGFR3, anti-Pdgfb anti-Dll4 and anti-CD34 antibody [85]. Gene expression of these proteins was found to be enriched in tip cells [83,86] and antibodies preferentially label endothelial tip cell bodies but less intensively their filopodia, endothelial stalk cells and phalanx cells. ESM1 is probably the most specific tip cell marker in mouse retinal tip cells, labeling additionally only some arterial ECs, whereas labeling of CXCR4 is high in tip cells, but also to a lesser extend present in some ECs of the vessel plexus, in arterial ECs and in perivascular cells [87]. The actin cytoskeleton provides a driving force for tip cell movement during angiogenesis, and labeling F-actin with phalloidin [83] is another way of identifying endothelial tip cells, especially by highlighting their filopodia. As an indirect method to distinguish tip cells from other EC phenotypes, labeling with antibodies against the adherens junction protein VE-cadherin, the adhesion protein PECAM1/CD31 or against proteins in the basal lamina (fibronectin, collagen, laminin) are used in combination with IB4, in which all ECs are labelled, but tip cell filopodia exclusively with IB4 [83,86]. The use of markers of the basal lamina is based on the concept that formation of new sprouts requires degradation of extracellular matrix to allow migration of tip cells, thus showing reduced staining in proximity of tip cells. Indeed, it was recently shown by triple-labeling of tip cells with F-actin, cortactin and collagen type IV that tip cells use so-called podosomes to degrade the extracellular matrix [88]. Labeling with anti-Ki67 or BrdU [83,86] is used as marker of proliferation, a property that is greatly reduced in tip cells.
A useful marker of tip cells employing in situ hybridization (ISH) is the Pdgfb gene [83,86]. Microarray analysis comparing DLL4+/- and wild-type mouse retinas [84] identified additional tip cell enriched genes, including,
apelin , angiopoietin-2 , chemokine receptor type 4 (CXCR4) and endocan ()[84,87,85], that these researchers confirmed to be tip cell specific by ISH and the latter two recently also by immunofluorescent staining [87].
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